1. Field of the Invention
The present invention relates to the identification of solvent-accessible amide hydrogens in polypeptides or proteins. The methods of the invention can be used to characterize the binding site involved in binding between a binding protein and a binding partner, and to study other changes in a polypeptide or protein which alter the rates at which hydrogen atoms exchange with solvent hydrogens, such as folding phenomena and other structural changes.
2. Background Art
Limitations of Current Methods of Characterizing Protein Binding Sites
Considerable experimental work and time are required to precisely characterize a binding site. In general, the techniques which are the easiest to use and which give the quickest answers, result in an inexact and only approximate idea of the nature of the critical structural features. Techniques in this category include the study of proteolytically generated fragments of the protein which retain binding function; recombinant DNA techniques, in which proteins are constructed with altered amino acid sequence (site directed mutagenesis); epitope scanning peptide studies (construction of a large number of small peptides representing subregions of the intact protein followed by study of the ability of the peptides to inhibit binding of the ligand to receptor); covalent crosslinking of the protein to its binding partner in the area of the binding site, followed by fragmentation of the protein and identification of crosslinked fragments; and affinity labeling of regions of the receptor which are located near the ligand binding site of the receptor, followed by characterization of such xe2x80x9cnearest neighborxe2x80x9d peptides. (Reviewed in 1, 2).
These techniques work best for the determination of the structure of binding subregions which are simple in nature, as when a single short contiguous stretch of polypeptide within a protein is responsible for most of the binding activity. However, for many protein-binding partner systems of current interest, the structures responsible for binding on both receptor and ligand or antibody are created by the complex interaction of multiple non-contiguous peptide sequences. The complexities of these interactions may confound conventional analytical techniques, as binding function is often lost as soon as one of the 3-dimensional conformations of the several contributing polypeptide sequences is directly or indirectly perturbed.
The most definitive techniques for the characterization of the structure of receptor binding sites have been NMR spectroscopy and X-ray crystallography. While these techniques can ideally provide a precise characterization of the relevant structural features, they have major limitations, including inordinate amounts of time required for study, inability to study large proteins, and, for X-ray analysis, the need for protein-binding partner crystals (Ref. 3).
Applicant""s technology overcomes these limitations and allows the rapid identification of each of the specific polypeptides and amino acids within a protein which constitute its protein ligand binding site or antibody binding subregion in virtually any protein-ligand system or protein antigen-antibody system, regardless of the complexity of the binding sites present or the size of the proteins involved. This technology is superior in speed and resolution to currently employed biochemical techniques.
Hydrogen (Proton) Exchange
When a protein in its native folded state is incubated in buffers containing heavy hydrogen (tritium or deuterium) labeled water, heavy hydrogen in the buffer reversibly exchanges with normal hydrogen present in the protein at acidic positions (for example, Oxe2x80x94H, Sxe2x80x94H, and Nxe2x80x94H groups) with rates of exchange which are dependent on each exchangeable hydrogen""s chemical environment, temperature, and most importantly, its accessibility to the tritiated water in the buffer. (Refs. 4, 5) Accessibility is determined in turn by both the surface (solvent-exposed) disposition of the hydrogen, and the degree to which it is hydrogen-bonded to other regions of the folded protein. Simply stated, acidic hydrogen present on amino acid residues which are on the outside (buffer-exposed) surface of the protein and which are hydrogen-bonded to solvent water will exchange more rapidly with heavy hydrogen in the buffer than will similar acidic hydrogen which are buried and hydrogen bonded within the folded protein.
Hydrogen exchange reactions can be greatly accelerated by both acid and base-mediated catalysis, and the rate of exchange observed at any particular pH is the sum of both acid and base mediated mechanisms. For many acidic hydrogen, a pH of 2.7 results in an overall minimum rate of exchange (Ref. 6, pg. 238, FIGS. 3a-c, refs. 7-11). While hydrogens in protein hydroxyl and amino groups exchange with tritium in buffer at millisecond rates, the exchange rate of one particular acidic hydrogen, the peptide amide bond hydrogen, is considerably slower, having a half life of exchange (when freely hydrogen bonded to solvent water) of approximately 0.5 seconds at 0xc2x0 C., pH 7, which is greatly slowed to a half life of exchange of 70 minutes at 0xc2x0 C. pH 2.7.
When peptide amide hydrogens are buried within a folded protein, or are hydrogen bonded to other parts of the protein, exchange half lives with solvent hydrogens are often considerably lengthened, at times being measured in hours to days. Hydrogen exchange at peptide amides is a fully reversible reaction, and rates of on-exchange (solvent heavy hydrogen replacing protein-bound normal hydrogen) are identical to rates of off-exchange (hydrogen replacing protein-bound heavy hydrogen) if the state of a particular peptide amide within a protein, including its chemical environment and accessibility to solvent hydrogens, remains identical during on-exchange and off-exchange conditions.
Hydrogen exchange is commonly measured by performing studies with proteins and aqueous buffers that are differentially tagged with pairs of the three isotopic forms of hydrogen (1H;Normal Hydrogen; 2H;Deuterium; 3H;Tritium). If the pair of normal hydrogen and tritium are employed, it is referred to as tritium exchange; if normal hydrogen and deuterium are employed, as deuterium exchange. Different physicochemical techniques are in general used to follow the distribution of the two isotopes in deuterium versus tritium exchange.
Tritium Exchange Techniques
Tritium exchange techniques (where the amount of the isotope is determined by radioactivity measurements) have been extensively used for the measurement of peptide amide exchange rates within an individual protein (reviewed in 4). The rates of exchange of other acidic protons (OH, NH, SH) are so rapid that they cannot be followed in these techniques and all subsequent discussion refers exclusively to peptide amide proton exchange. In these studies, purified proteins are on-exchanged by incubation in buffers containing tritiated water for varying periods of time, transferred to buffers free of tritium, and the rate of off-exchange of tritium determined. By analysis of the rates of tritium on- and off-exchange, estimates of the numbers of peptide amide protons in the protein whose exchange rates fall within particular exchange rate ranges can be made. These studies do not allow a determination of the identity (location within the protein""s primary amino acid sequence) of the exchanging amide hydrogens measured.
Extensions of these techniques have been used to detect the presence within proteins of peptide amides which experience allosterically-induced changes in their local chemical environment and to study pathways of protein folding (5, 12-14). For these studies, tritium on-exchanged proteins are allowed to off-exchange after they have experienced either an allosteric change in shape, or have undergone time-dependent folding upon themselves, and the number of peptide amides which experience a change in their exchange rate subsequent to the allosteric/folding modifications determined. Changes in exchange rate indicate that alterations of the chemical environment of particular peptide amides have occurred which are relevant to proton exchange (solvent accessibility, hydrogen bonding etc.). Peptide amides which undergo an induced slowing in their exchange rate are referred to as xe2x80x9cslowed amidesxe2x80x9d and if previously on-exchanged tritium is sufficiently slowed in its off-exchange from such amides there results a xe2x80x9cfunctional tritium labelingxe2x80x9d of these amides. From these measurements, inferences are made as to the structural nature of the shape changes which occurred within the isolated protein. Again, determination of the identity of the particular peptide amides experiencing changes in their environment is not possible with these techniques.
Four groups of investigators have described technical extensions (collectively referred to as medium resolution tritium exchange) which allow the locations of particular slowed, tritium labeled peptide amides within the primary sequence of small proteins to be localized to a particular proteolytic fragment, though not to a particular amino acid.
Rosa and Richards were the first to describe and utilize medium resolution tritium techniques in their studies of the folding of ribonuclease S protein fragments (15-17). However, the techniques described by Rosa and Richards were of marginal utility, primarily due to their failure to optimize certain critical experimental steps (reviewed in 6, pg 238, 244). No studies employing related techniques were published until the work of Englander and co-workers in which extensive modifications and optimizations of the Rosa and Richards technique were first described.
Englander""s investigations utilizing tritium exchange have focused exclusively on the study of allosteric changes which take place in tetrameric hemoglobin (a subunit and b subunit 16 kD in size each) upon deoxygenation (6,18-21). In the Englander procedure, native hemoglobin (milligram quantities) in the oxygenated state is on-exchanged in tritiated water of relatively low specific activity (2-100 mCi/ml). The hemoglobin is then deoxygenated (inducing allosteric change), transferred to tritium-free buffers by gel permeation column chromatography, and then allowed to out-exchange for 10-50 times the on-exchange time. On-exchanged tritium present on peptide amides which experience no change in exchange rate subsequent to the induced allosteric change in hemoglobin structure off-exchanges at rates identical to its on-exchange rates, and therefore is almost totally removed from the protein after the long off-exchange period. However, peptide amides which experience slowing of their exchange rate subsequent to the induced allosteric changes preferentially retain the tritium label during the period of off-exchange.
To localize (in terms of hemoglobin""s primary sequence) the slowed amides bearing the residual tritium label, Englander then proteolytically fragments the off-exchanged hemoglobin with the protease pepsin, separates, isolates and identifies the various peptide fragments by reverse phase high pressure liquid chromatography (RP-HPLC), and determines which fragments bear the residual tritium label by scintillation counting. However, as the fragmentation of hemoglobin proceeds, each fragment""s secondary and tertiary structure is lost and the unfolded peptide amides become freely accessible to H2O in the buffer. At physiologic pH ( greater than 6), any amide-bound tritium label would leave the unfolded fragments within seconds. Englander therefore performs the fragmentation and HPLC peptide isolation procedures under conditions which he believes minimize peptide amide proton exchange, including cold temperature (4xc2x0 C.) and use of phosphate buffers at pH 2.7 (reviewed in 6). This technique has been used successfully by Englander to coarsely identify and localize the peptidic regions of hemoglobin xcex1 and xcex2 chains which participate in deoxygenation-induced allosteric changes (18-21). The ability of the Englander technique to localize tritium labeled amides, while an important advance, remains low; at the best, Englander reports that his technique localizes amide tritium label to hemoglobin peptides 14 amino acids or greater in size, without the ability to further sublocalize the label.
Moreover, in Englander""s work, there is no appreciation that a suitably adapted tritium exchange technique might be used to identify the peptide amides which reside in the contacting surface of a protein receptor and its binding partner: his disclosures are concerned exclusively with the mapping of allosteric changes in hemoglobin. Furthermore, based on his optimization studies (6-11,13), Englander teaches and warns that a pH of 2.7 must be employed in both the proteolysis and HPLC steps, necessitating the use of proteases which are functional at these pH""s (acid proteases). Unfortunately, acid proteases are relatively nonspecific in their sites of proteolytic cleavage, leading to the production of a very large number of different peptide fragments and hence to considerable HPLC separation difficulties. The constraint of performing the HPLC separation step at pH 2.7 greatly limits the ability to optimize the chromatographic separation of multiple overlapping peptides by varying the pH at which HPLC is performed. Englander tried to work around these problems, for the localization of hemoglobin peptides experiencing allosteric changes, by taking advantage of the fact that some peptide bonds are somewhat more sensitive to pepsin than others. He therefore limits the duration of exposure of the protein to pepsin to reduce the number of fragments. Even then the fragments were xe2x80x9cdifficult to separate cleanlyxe2x80x9d. They were also, of course, longer (on average), and therefore the resolution was lower. He also tried to simplify the patterns by first separating the alpha and beta chains of hemoglobin. However, there was a tradeoff: increased tritium loss during the alpha-beta separation and the removal of the solvent, preparatory to proteolysis. Englander concludes,
xe2x80x83xe2x80x9cAt present the total analysis of the HX (hydrogen exchange) behavior of a given protein by these methods is an immense task. In a large sense, the best strategies for undertaking such a task remain to be formulated. Also, these efforts would benefit from further technical improvements, for example in HPLC separation capability and perhaps especially in the development of additional acid proteases with properties adapted to the needs of these experimentsxe2x80x9d (6).
Over the succeeding years since this observation was made, no advances have been disclosed which address these critical limitations of the medium resolution tritium exchange technique. It has been perceived that improvements to the HPLC separation step were problematic due to the constraint of working at pH 2.7. The current limited success with small proteins has made it pointless to attempt similar studies of larger proteins where the problems of inadequate HPLC peptide separation at pH 2.7, and imprecision in the ability to sublocalize labeled amides would be greatly compounded. Furthermore, most acid-reactive proteases are in general no more specific in their cleavage patterns than pepsin and efforts to improve the technology by employing other acid reactive proteases other than pepsin have not significantly improved the technique. Given these limitations of medium resolution tritium exchange art, no studies have been disclosed which utilize proteins with subunit size greater than 16 kilodaltons.
Allewell and co-workers have disclosed studies utilizing the Englander techniques to localize induced allosteric changes in the enzyme Escherichia coli aspartate transcarbamylase (22,23). Burz, et al. (22) is a brief disclosure in which the isolated R2 subunit of this enzyme is on-exchanged in tritiated buffer of specific activity 100 mCi/ml, allosteric change induced by the addition of ATP, and then the conformationally altered subunit off-exchanged. The enzyme R2 subunit was then proteolytically cleaved with pepsin and analyzed for the amount of label present in certain fragments. Analysis employed techniques which rigidly adhered to the recommendations of Englander, utilizing a single RP HPLC separation in a pH 2.8 buffer.
The authors note difficulty in separating the large number of peptides generated, even from this small protein subfragment, given the constraints of the Englander methodology. They comment that xe2x80x9cthe principal limitation of this method at present is the separation with columns now availablexe2x80x9d. ATP binding to the enzyme was shown to alter the rate of exchange of hydrogens within several relatively large peptidic fragments of the R2 subunit. In a subsequent more complete disclosure (23), the Allewell group discloses studies of the allosteric changes induced in the R2 subunit by both ATP and CTP. They disclose on-exchange of the R2 subunit in tritiated water-containing buffer of specific activity 22-45 mCi/ml, addition of ATP or CTP followed by off exchange of the tritium in normal water-containing buffer. The analysis comprised digestion of the complex with pepsin, and separation of the peptide fragments by reverse phase HPLC in a pH 2.8 or pH 2.7 buffer, all of which rigidly adheres to the teachings of Englander. Peptides were identified by amino acid composition or by N-terminal analysis, and the radioactivity of each fragment was determined by scintillation counting. In both of these studies the localization of tritium label was limited to peptides which averaged 10-15 amino acids in size, without higher resolution being attempted.
Beasty, et al. (24) have disclosed studies employing tritium exchange techniques to study folding of the a subunit of E. Coli tryptophan synthetase. The authors employed tritiated water of specific activity 20 mCi/ml, and fragmented the tritium labeled enzyme protein with trypsin at a pH 5.5, conditions under which the protein and the large fragments generated retained sufficient folded structure as to protect amide hydrogens from off exchange during proteolysis and HPLC analysis. Under these conditions, the authors were able to produce only 3 protein fragments, the smallest being 70 amino acids in size. The authors made no further attempt to sublocalize the label by further digestion and/or HPLC analysis. Indeed, under the experimental conditions they employed (they performed all steps at 12xc2x0 C. instead of 4xc2x0 C., and performed proteolysis at pH 5.5 instead of pH in the range of 2-3), it would have been impossible to further sublocalize the labeled amides by tritium exchange, as label would have been immediately lost (off-exchanged) by the unfolding of subsequently generated proteolytic fragments at pH 5.5 if they were less than 10-30 amino acids in size.
Fromageot, et al., U.S. Pat. No. 3,828,102 (25) discloses using hydrogen exchange to tritium label a protein and its binding partner, and Benson, U.S. Pat. No. 3,560,158 and 3,623,840 (26) disclose using hydrogen exchange to tritiate compounds for analytical purposes.
However, none of the methods described in the art are capable of localizing the positions of the tritium labels of the labeled proteins at high resolution, the best resolution in the art generally being on the order of xe2x89xa714 amino acid residues.
Deuterium Exchange Techniques
Fesik, et al (27) discloses measuring by NMR the hydrogen (deuterium) exchange of a peptide before and after it is bound to a protein. From this data, the interactions of various hydrogens in the peptide with the binding site of the protein are analyzed.
Patterson, et al. (28) and Mayne, et al. (29) disclose NMR mapping of an antibody binding site on a protein (cytochrome-C) using deuterium exchange. This relatively small protein, with a solved NMR structure, is first complexed to anti-cytochrome-C monoclonal antibody, and the preformed complex then incubated in deuterated water-containing buffers and NMR spectra obtained at several time intervals. The NMR spectra of the antigen-antibody complex is examined for the presence of peptide amides which experience slowed hydrogen exchange with solvent deuterium as compared to their rate of exchange in uncomplexed native cytochrome-C. Benjamin, et al. (30) employ an identical NMR-deuterium technique to study the interaction of hen egg lysozyme (HEL) with HEL-specific monoclonal antibodies. While both this NMR-deuterium technique, and medium resolution tritium exchange rely on the phenomenon of proton exchange at peptide amides, they utilize radically different methodologies to measure and localize the exchanging amides. Furthermore, study of proteins by the NMR technique is not possible unless the protein is small (less than 30 kD), large amounts of the protein are available for the study, and computationally intensive resonance assignment work is completed.
Recently, others (45-50) have disclosed techniques in which exchange-deuterated proteins are incubated with binding partner, off-exchanged, the complex fragmented with pepsin, and deuterium-bearing peptides identified by single stage fast atom bombardment (Fab) or electrospray mass spectroscopy (MS). In these studies, no attempt has been made to sublocalize peptide-bound deuterium within the proleolytically or otherwise generated peptide fragments.
Thus, as is evidenced by the above discussion, there remains a need in the art for simple and efficient methods whereby the positions of labeled solvent-accessible peptide amide hydrogens can be localized at high resolution within the primary amino acid sequence of a polypeptide or protein, as well as simple and efficient methods for studying or mapping the binding sites and/or interaction surfaces of a polypeptide or protein. Accordingly, these are objects of the present invention.
These and other shortcomings in the art are overcome by the present invention, which in one aspect provides methods for the functional labeling and identification of specific amino acid residues that participate in binding protein-binding partner interactions. The methods of the invention are particularly suitable for the study of the binding protein-binding partner subregions of large ( greater than 30 KD) proteins, even in small quantities.
In one embodiment, the label is tritium and the amount of label on a fragment or subfragment is determined by measuring its radioactivity. In a second embodiment, the label is deuterium and the amount of label on a fragment or subfragment is determined by mass spectrometry. The term xe2x80x9cheavy hydrogenxe2x80x9d is used herein to refer generically to either tritium or deuterium. In addition, references to tritium apply mutatis mutandis to deuterium except when clearly excluded.
In essence, the binding protein is first tritiated or deuterated under conditions wherein native hydrogens are replaced by the tritium or deuterium label (this is the xe2x80x9con-exchangexe2x80x9d step). Then the binding partner is allowed to interact with labeled protein. The binding partner occludes the binding site and protects the tritium or deuterium labels of that site from a subsequent xe2x80x9coff-exchangexe2x80x9d. Thus, after the xe2x80x9coff-exchangexe2x80x9d, only the binding site residues are labeled. Since the binding site is normally only a small portion of the molecules, a higher signal-to-background ratio is obtained with this approach than with Englander""s more conventional procedure.
In order to actually identify the labeled residues, one must first dissociate the complex under slow hydrogen isotope exchange (H3/H1 or H2/H1) conditions, since otherwise the labels would leave the binding site as soon as the ligand was removed. The binding protein is then optionally fragmented (e.g., with an endoprotease such as pepsin), still under slow hydrogen exchange conditions, to obtain fragments. Those fragments which bear label presumably include binding site residues. At this point, the resolution of the binding site is no better than the fragment size.
A finer localization of the labels is achieved by analysis of subfragments generated by controlled, stepwise, degradation of the binding protein or of each isolated, labeled peptide fragment (if the binding protein was optionally fragmented) under slowed exchange conditions. For the purpose of the present invention, the protein or a peptide fragment is said to be xe2x80x9cprogressivelyxe2x80x9d, xe2x80x9cstepwisexe2x80x9d or xe2x80x9csequentiallyxe2x80x9d degraded if a series of fragments are obtained which are similar to those which would be achieved by an ideal exopeptidase. For an ideal exopeptidase, only an end amino acid is removed. Thus, if the n amino acids of a peptide were labeled A1 to An (the numbering starting at whichever end the degradation begins), the series of subfragments produced by an ideal exopeptidase would be A2 . . . An, A3 . . . An, . . . , Anxe2x88x921xe2x88x92An, and finally An. However, it is to be understood that while preferably each subfragment of the series of subfragments obtained is shorter than the preceding subfragment in the series by a single terminal amino acid residue, exopeptidases are not necessarily ideal. Thus, for purposes of the present invention, a fragment is said to be xe2x80x9cprogressively,xe2x80x9d xe2x80x9cstepwisexe2x80x9d or xe2x80x9csequentiallyxe2x80x9d degraded if a series of subfragments is generated wherein each subfragment in the series is composed of about 1-5 fewer terminal amino acid residues than the preceding subfragment in the series. The signals produced by the successive subfragments are correlated in order to determine which amino acids of the fragment in question were labeled.
This procedure was not used in any of the cited references to further localize the labeling sites, though improved resolution was certainly a goal of the art. The closest the art comes is Englander""s general suggestions of further fragmentations with another xe2x80x9cacid proteasexe2x80x9d.
The progressive degradation is preferably achieved by an enzyme, and more preferably by a carboxypeptidase. The need to employ an acidic pH at the time of degradation to minimize tritium losses discourages use of carboxypeptidases which are substantially inactivated by the required acidic buffers. However, carboxypeptidase-P, carboxypeptidase Y, and several other acid-reactive (i.e., enzymatically active under acid conditions) carboxypeptidases are suitable for proteolysis of peptides under acidic conditions, even at pH 2.7.
Progressive subfragmentation of purified tritium label-bearing peptides is performed with acid-reactive carboxypeptidases under conditions that produce a complete set of amide-labeled daughter peptides each shorter than the preceding one by 1-5 carboxy terminal amino acids, and preferably by a single carboxy-terminal amino acid. HPLC analysis of the several members of this set of progressively truncated peptides allows the reliable assignment of label to a particular amide position within the parent peptide.
Alternatively, the present invention contemplates C-terminal chemical degradation techniques that can be performed under xe2x80x9cslow hydrogen exchange conditionsxe2x80x9d e.g., by pentafluoropropionic acid anhydride. The sensitivity of the technique may be improved by the use of reference peptide subfragments as HPLC mobility markers.
In general, the art has given insufficient consideration to the problems of denaturing the binding protein sufficiently to facilitate proteolysis under slow hydrogen exchange conditions. Pepsin, for example, is much less active at 0xc2x0 C. than at room temperature. While pepsin is able to extensively digest hemoglobin that has been denatured by acidic pH at 0xc2x0 C., certain other binding proteins, such as hen egg lysozyme, are much more resistant to denaturation by slow H-exchange conditions, and hence to subsequent pepsin digestion. As a result, many fewer and longer fragments are generated. This complicates the analysis.
In a preferred embodiment, the labeled binding protein is exposed, before fragmentation, to denaturing conditions compatible with slow hydrogen exchange and sufficiently strong to denature the protein enough to render it adequately susceptible to the intended proteolytic treatment. If these denaturing conditions would also denature the protease, then, prior to proteolysis, the denatured protein is switched to less denatured conditions (still compatible with slow H-exchange) sufficiently denaturing to maintain the protein in a protease-susceptible state but substantially less harmful to the protease in question.
Preferably, the initial denaturant is guanidine thiocyanate, and the less denaturing condition is obtained by dilution with guanidine HCl.
Disulfide bonds, if present in the binding protein to be digested, can also interfere with analysis. Disulfide bonds can hold the protein in a folded state where only a relatively small number of peptide bonds are exposed to proteolytic attack. Even if some peptide bonds are cleaved, failing to disrupt the disulfide bonds would reduce resolution of the peptide fragments still joined to each other by the disulfide bond; instead of being separated, they would remain together. This would reduce the resolution by at least a factor of two (possibly more, depending on the relationship of disulfide bond topology to peptide cleavage sites). If the disulfide bonds are not disrupted, further sublocalization of the tritium-labeled amides within each of the disulfide-joined peptides would be very difficult, as amino acid removal would occur, at different times and at different rates, at each C-terminal of the disulfide linked segments.
The applicant has discovered that water soluble phosphines may be used to disrupt a protein""s disulfide bonds under xe2x80x9cslow hydrogen exchangexe2x80x9d conditions. This allows much more effective fragmentation of large proteins which contain disulfide bonds without causing tritium label to be lost from the protein or its proteolytic fragments (as would be the case with conventional disulfide reduction techniques which must be performed at pH""s which are very unfavorable for preservation of tritium label).
In another embodiment, peptide amides on the binding protein""s surface are indirectly labeled by transfer of tritium or deuterium that has been previously attached by hydrogen exchange to the interaction surface of the binding partner. This procedure will functionally label receptor protein amides if they are slowed by complex formation and are also in intimate contact with the binding partner, in the complexed state. Amides that are distant from the interaction surface but slowed in exchange because of complex formation-induced allosteric changes in the protein will not be labeled.
In another aspect, the present invention provides a method for determining which peptide amide hydrogens in a polypeptide or protein are accessible to solvent. By using the method of the invention, the positions of solvent-accessible peptide amide hydrogens within a polypeptide or protein can be localized at high resolution, i.e., typically within 5 or fewer amino acid residues, and in many instances to within a single amino acid residue.
In the method, solvent accessible peptide amide hydrogens of a polypeptide or protein of interest are on-exchanged by contacting the polypeptide or protein with heavy hydrogen under conditions wherein the native solvent-accessible peptide amide hydrogens are replaced with heavy hydrogen (deuterium or tritium), such as, for example, physiological conditions wherein the polypeptide or protein is folded into its native conformation. Peptide amide protons that are inaccessible to solvent, such as those that are buried within the interior of the polypeptide or protein structure or those that participate in intramolecular hydrogen-bonding interactions, do not readily exchange with the heavy hydrogens in the solvent. Thus, those peptide amide hydrogens that are solvent-accessible are selectively labeled with heavy hydrogen.
The positions of the labeled peptide amide hydrogens within the polypeptide or protein can be localized at high resolution by progressively generating a series of subfragments under conditions of slowed exchange as previously described, determining which subfragments are labeled and correlating the sequences of the labeled subfragments with the sequence of the polypeptide or protein to determine which peptide amide groups in the polypeptide or protein were labeled, and thus accessible to solvent.
In some embodiments, especially those wherein the polypeptide or protein of interest is relatively large, the polypeptide or protein may optionally be first fragmented (e.g., with an endoprotease or mixture of endoproteases) under conditions of slowed exchange as previously described, and the positions of the labels localized to high resolution by progressively degrading each labeled fragment into a series of subfragments, determining which subfragments are labeled and correlating the sequences of the labeled subfragments to the sequences of the labeled fragments and, ultimately, to the sequence of the polypeptide or protein, as previously described.
In a preferred embodiment, the polypeptide or protein is denatured and any disulfide bonds reduced under conditions of slowed exchange prior to fragmentation and/or subfragmentation.